Insects and other pests cost farmers billions of dollars annually in crop losses and expense to keep these pests under control. In addition to losses in field crops, insect pests are also a burden to vegetable and fruit growers, to producers of ornamental flowers, and to home gardeners. The losses caused by insect pests in agricultural production environments include decrease in crop yield, reduced crop quality, and increased harvesting costs.
Insect pests are mainly controlled by intensive applications of chemical pesticides, which are active through inhibition of insect growth, prevention of insect feeding or reproduction, or cause death. Good insect control can thus be reached, but these chemicals can sometimes affect other beneficial insects. Another problem resulting from the wide use of chemical pesticides is the appearance of resistant insect populations. This has been partially alleviated by various resistance management practices, but there is an increasing need for alternative pest control agents. Biological pest control agents, such as Bacillus thuringiensis (B.t.) strains expressing pesticidal toxins like delta-endotoxins, have also been applied to crop plants with satisfactory results, offering an alternative or compliment to chemical pesticides. The genes coding for some of these delta-endotoxins have been isolated and their expression in heterologous hosts have been shown to provide another tool for the control of economically important insect pests. In particular, the expression of insecticidal toxins, such as Bacillus thuringiensis delta-endotoxins, in transgenic plants have provided efficient protection against selected insect pests, and transgenic plants expressing such toxins have been commercialized, allowing farmers to reduce applications of chemical insect control agents.
Lepidopterans are an important group of agricultural, horticultural, and household pests which cause a large amount of damage each year. This insect order encompasses foliar- and root-feeding larvae and adults. Lepidopteran insect pests include, but are not limited to: Achoroia grisella, Acleris gloverana, Acleris variana, Adoxophyes orana, Agrotis ipsilon (black cutworm “BCW”), Alabama argillacea, Alsophila pometaria, Amyelois transitella, Anagasta kuehniella, Anarsia lineatella, Anisota senatoria, Antheraea pernyi, Anticarsia gemmatalis (velvetbean caterpillar “VBC”), Archips sp., Argyrotaenia sp., Athetis mindara, Bombyx mori, Bucculatrix thurberiella, Cadra cautella, Choristoneura sp., Cochylls hospes, Colias eurytheme, Corcyra cephalonica, Cydia latiferreanus, Cydia pomonella, Datana integerrima, Dendrolimus sibericus, Desmia feneralis, Diaphania hyalinata, Diaphania nitidalis, Diatraea grandiosella (southwestern corn borer “SWCB”), Diatraea saccharalis, Ennomos subsignaria, Eoreuma loftini, Esphestia elutella, Erannis tilaria, Estigmene acrea, Eulia salubricola, Eupocoellia ambiguella, Eupoecilia ambiguella, Euproctis chrysorrhoea, Euxoa messoria, Galleria mellonella, Grapholita molesta, Harrisina americana, Helicoverpa subflexa, Helicoverpa zea (corn earworm “CEW”), Heliothis virescens (tobacco budworm “TBW”), Hemileuca oliviae, Homoeosoma electellum, Hyphantia cunea, Keiferia lycopersicella, Lambdina fiscellaria fiscellaria, Lambdina fiscellaria lugubrosa, Leucoma salicis, Lobesia botrana, Loxostege sticticalis, Lymantria dispar, Macalla thyrisalis, Malacosoma sp., Mamestra brassicae, Mamestra configurata, Manduca quinquemaculata, Manduca sexta, Maruca testulalis, Melanchra picta, Operophtera brumata, Orgyia sp., Ostrinia nubilalis (European corn borer “ECB”), Paleacrita vernata, Papiapema nebris (common stalk borer), Papilio cresphontes, Pectinophora gossypiella, Phryganidia californica, Phyllonorycter blancardella, Pieris napi, Pieris rapae, Plathypena scabra, Platynota flouendana, Platynota stultana, Platyptilia carduidactyla, Plodia interpunctella, Plutella xylostella (diamondback moth “DBM”), Pontia protodice, Pseudaletia unipuncta, Pseudoplusia includens (soybean looper “SBL”), Sabulodes aegrotata, Schizura concinna, Sitotroga cerealella, Spilonta ocellana, Spodoptera eridania (southern armyworm “SAW”), Spodoptera frugiperda (fall armyworm “FAW”), Spodoptera exigua (beet armyworm “BAW”), Thaurnstopoea pityocampa, Ensola bisselliella, Trichoplusia ni (cabbage looper “CL”), Udea rubigalis, Xylomyges curiails, and Yponomeuta padella. Any genus listed above (and others), generally, can also be targeted as a part of the subject invention. Any additional insects in any of these genera (as targets) are also included within the scope of this invention.
Bacillus thuringiensis (B.t.) is a soil-borne, Gram-positive, spore forming bacterium that produces insecticidal crystal proteins known as delta endotoxins or Cry proteins (reviewed in Schnepf et al., 1998). Novel Crystal (Cry) proteins with new insecticidal properties continue to be discovered at an increasing rate, and over 440 Cry genes have been reported. Currently, there are over 450 unique Cry and Cytotoxin (Cyt) proteins classified among 57 primary homology ranks. Cry proteins are named based on the degree of sequence identity, with primary, secondary and tertiary boundaries occurring at approximately 45%, 78% and 95% identity, respectively; close alleles are assigned new quaternary designations (Crickmore et al., 1998). An expansive list of delta endotoxins is maintained and regularly updated at lifesci.sussex.ac.uk/home/Neil_Crickmore/Bt/intro.html. There are currently over 73 main groups of “Cry” toxins (Cry1-Cry73), with additional Cyt toxins and Vegetative Insecticidal Protein (VIP) toxins and the like. Many of each numeric group have capital-letter subgroups, and the capital letter subgroups have lower-cased letter sub-subgroups. (Cry1 has A-L, and Cry1A has a-i, for example).
B.t. proteins have been used to create the insect-resistant transgenic plants that have been successfully registered or deregulated and commercialized to date. These include Cry1Ab, Cry1Ac, Cry1F, Vip3A, Cry34Ab1/Cry35Ab1, and Cry3Bb in corn, Cry1Ac, Vip3A and Cry2Ab in cotton, and Cry3A in potato. B.t. toxins represent over 90% of the bioinsecticide market and essentially the entire source of genes for transgenic crops that have been developed to provide resistance to insect feeding.
Cry proteins are oral intoxicants that function by acting on midgut cells of susceptible insects. The active forms of many Cry proteins comprise three distinct protein domains. The most well studied B.t. proteins are members of the three-domain Cry delta-endotoxins. These proteins range in size from approximately 70 kDa to 130 kDa. Primary protein sequence analysis reveals five highly conserved sequence blocks and a high degree of sequence variability between conserved blocks three and five (Schnepf et al., 1998).
Three dimensional crystal structures have been determined for Cry1Aa1, Cry2Aa1, Cry3Aa1, Cry3Bb1, Cry4Aa, Cry4Ba and Cry8Ea1 as examples. These structures are remarkably similar and are comprised of three distinct domains with the following features (reviewed in de Maagd et al., 2003). Domain I is a bundle of seven alpha helices where helix five is surrounded by six amphipathic helices. This domain has been implicated in midgut membrane insertion and pore formation. It shares homology with other pore forming proteins including hemolysins and colicins. Domain II is comprised of three anti-parallel beta sheets packed together in a beta prism. This domain shares homology with certain carbohydrate-binding proteins including vitelline and jacaline. The loops of this domain play important roles in binding insect midgut receptors. In Cry1A proteins, surface exposed loops at the apices of domain II beta sheets are involved in binding to lepidopteran cadherin receptors. Domain III is a beta sandwich structure that interacts with a second class of receptors, examples of which are aminopeptidase and alkaline phosphatase in the case of Cry1A proteins (Piggot and Ellar, 2007). Structurally this domain is related to carbohydrate-binding domains of proteins such as glucanases, galactose oxidase, sialidase and others. This domain binds certain classes of receptor proteins and perhaps participates in insertion of an oligomeric toxin pre-pore. Conserved B.t. sequence blocks 2 and 3 map near the N-terminus and C-terminus of domain 2, respectively. Hence, these conserved sequence blocks 2 and 3 are approximate boundary regions between the three functional domains. These regions of conserved DNA and protein homology have been exploited for engineering recombinant B.t. toxins (U.S. Pat. No. 6,090,931, WO 91/01087, WO95/06730, WO 1998022595).
One proposed model for Cry protein mode of action is based on pore formation in the midgut membranes of susceptible insects (Knowles and Ellar, 1987). In the current version of this model (Bravo et al., 2007), binding to both cadherin and aminopeptidase receptors on Lepidopteran midgut membranes are required for Cry protein toxicity. According to the pore formation model, Cry protein intoxication involves several steps: 1) Proteolytic processing of soluble Cry protoxin to an activated core toxin; 2) Cry protein binding to cadherin receptors on the insect midgut; 3) further proteolytic cleavage at the core toxin N-terminus to remove an α-helical region; 4) Cry protein oligomerization to form a pre-pore; 5) pre-pore binding to second site membrane receptors (aminopeptidases and alkaline phosphatases); 6) pre-pore insertion into the membrane and 7) osmotic cell lysis leading to midgut disruption and insect death.
The widespread adoption of insect-resistant transgenic plant technology gives rise to a concern that pest populations will develop resistance to the insecticidal proteins produced by these plants. Several strategies have been suggested for preserving the utility of B.t.-based insect resistance traits which include deploying proteins at a high dose in combination with a refuge, and alternation with, or co-deployment of, different toxins (McGaughey et al. (1998), “B.t. Resistance Management,” Nature Biotechnol. 16:144-146).
The development of insect resistance to B.t. Cry proteins can result through several mechanisms (Heckel et al., 2007, Piggot and Ellar, 2007). Multiple receptor protein classes for Cry proteins have been identified within insects, and multiple examples exist within each receptor class. Resistance to a particular Cry protein may develop, for example, by means of a mutation within the toxin-binding portion of a cadherin domain of a receptor protein. A further means of resistance may be mediated through a protoxin-processing protease. Thus, resistance to Cry1A toxins in species of Lepidoptera has a complex genetic basis, with at least four distinct, major resistance genes. Lepidopteran insects resistant to Cry proteins have developed in the field for Plutella xylostella (Tabashnik, 1994), Trichoplusia ni (Janmaat and Myers 2003, 2005), Helicoverpa zea (Tabashnik et al., 2008), and Spodoptera frupperda (Storer, et al., 2010). Development of new high potency Cry proteins will provide additional tools for management of Lepidopteran insect pests.
This invention provides B.t. insecticidal proteins that are effective in controlling insects that are resistant to Cry1Ac and Cry1F. These protein toxins may be used advantageously to protect agronomic crops from insect feeding damage. The ability to express these insect toxins in such a manner that sufficient quantity of the functionally active protein is present in a crop of interest is also a subject of this invention.